An unusual feruloyl esterase belonging to family VIII esterases and... substrate range

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An unusual feruloyl esterase belonging to family VIII esterases and... substrate range
An unusual feruloyl esterase belonging to family VIII esterases and displaying a broad
substrate range
Colin W Ohlhoffe, Bronwyn M Kirbya, Ana Casanuevab, Robert J Huddyc, Rolene Bauera,
David LR Mutepfad, Don A Cowana,d and Marla Tuffina
Institute for Microbial Biotechnology and Metagenomics, University of the Western Cape,
Bellville, Cape Town, South Africa
Technology Transfer Office, University of the Western Cape, Bellville, Cape Town, South
Centre for Bioprocess Engineering Research (CeBER), Department of Chemical
Engineering, University of Cape Town, Rondebosch, Cape Town, 7701, South Africa
Centre for Microbial Ecology and Genomics, University of Pretoria, Pretoria, South Africa
Fruitique (Pty) Ltd, 27 Scheckter Rd Killarney Gardens, Cape Town, 7441, South Africa
*[email protected], +27219599725, +27 21 9591506
A thermophilic compost metagenomic library constructed in E. coli was functionally
screened for novel esterases. Of the 110,592 fosmid clones screened, 25 clones demonstrated
degradative activity on glyceryl tributyrate (a hit rate of 1:4,423). Four clones displayed
ferulic acid esterase activity and were sequenced using 454 Titanium sequencing technology.
EstG34, a 410 amino acid protein, was identified as having high sequence identity with a
number of bacterial β-lactamases. EstG34 has the S-X-X-K motif which is conserved in class
C β-lactamases and Family VIII carboxylesterases. Purified recombinant Est34 had a
molecular mass of 42 kDa and displayed hydrolytic activity towards a variety of
p-nitrophenylesters, hydroxycinnamic acid esters and α-naphthol acetate. EstG34 represents
the first Family VIII carboxylesterase and β-lactamase fold enzyme, able to hydrolyse
ferulate and a number of other hydroxycinnamic acid esters. In addition, EstG34 is the first
reported FAE to not adopt the α/β hydrolase conformation. The sequence similarity and wide
substrate utilization capability of this esterase complicates its placement within current
classification systems, but also draws attention to the enzyme‟s potential versatility in a range
of industrial applications.
Metagenome, Alkaliphilic, Compost, Feruloyl Esterase, family VIII carboxylesterase
Microbial communities are a rich source of metabolic and genomic diversity, and are critical
components of functioning ecosystems. Both direct cultivation techniques and indirect
molecular methods have been used to investigate and exploit this wealth of microbial
diversity. It is now widely accepted that traditional methods involving the cultivation and
isolation of microorganisms only provide limited access to the total genetic information
within an environment [1]. Metagenomic methods entailing the isolation and cloning of
microbial DNA directly from environmental samples circumvent the limitations of culturing
and have been used to identify a variety of biotechnologically relevant microbial products [2,
Composting is a self-heating, aerobic, solid-phase biological system which accelerates the
natural process of biodegradation and mineralisation of organic matter. Microorganisms are
essential to the composting process and compost microbial communities have been shown to
produce a wide range of robust lignocellulolytic enzymes [4]. Carboxylic esterases are a
diverse class of hydrolases which catalyze the cleavage and formation of ester bonds,
including ester linkages in plant cell wall polysaccharides. They are commercially significant
biocatalysts in biotechnological processes, and have a wide range of industrial and medicinal
applications [5, 6]. Carboxylic esterases are ubiquitous in nature, having been identified in all
domains of life (Bacteria, Archaea and Eukaryotes), as well as in some viruses [7]. Within the
carboxylic ester hydrolase family, there are two recognized groups; the lipases (EC,
triacyglycerol hydrolases) and esterases (EC, carboxylester hydrolases). The current
sub-classification of esterases is based on biochemical and physiological properties of the
enzymes, particularly substrate specificities, as well as primary and tertiary structures [8].
Here we present the use of metagenomics to identify a novel Group VIII esterase with broad
feruloyl and aromatic ester substrate specificity.
Materials and Methods
Samples and Strains
E. coli strains EPI300 (Epicentre® Biotechnologies), GeneHogs (Invitrogen) and Rosetta
pLysS (Novagen) were used for all cloning and expression studies. Compost samples were
collected from a commercial compost production facility located in the Western Cape
Province of South Africa during the summer season of 2009 (GPS position 34°S 2‟ 53.35‟‟,
18°E 31‟ 45.71”). The compost source material consisted of an unspecified mix of wood
chips and sawdust, with lesser amounts of plant debris and bovine manure. The average
temperature of the compost at the point of sampling was 70 °C and the pH was 6.1. Elemental
analysis (BemLab Laboratories, Strand, South Africa) indicated the composition of the
compost was as follows: C 25.8%; N 1.1%; K 0.72%.
Metagenomic fosmid library construction
Metagenomic DNA was extracted using the chemical lysis method described by Zhou et al.
[9] with slight modifications (addition of SDS (2% w/v) and PVPP (0.5% w/v) to the
extraction buffer). Persistent humic- and phenolic compounds were removed by
electrophoresis in low melting point (LMP) agarose. DNA samples were loaded onto a 0.7%
LMP agarose gel in 1 X TAE buffer and electrophoresis was performed overnight at 1.5
V/cm. High-molecular weight (>35 kb) DNA fragments were excised from LMP agarose and
treated with agarase (Fermentas) followed by isopropanol precipitation according to the
manufacturer‟s instructions. DNA was quantitated using the Quanti-iT dsDNA BR assay kit
with a Qubit fluorimeter as described by the manufacturer (Invitrogen). The metagenomic
DNA library was constructed using the CopyControl™ Fosmid Library production kit
(Epicentre® Biotechnologies) according to the manufacturer‟s guidelines. Briefly, purified
metagenomic DNA was end-repaired and ligated to the CopyControl pCC1Fos™ vector.
Fosmid clones were packaged by MaxPlax Lambda phage and transfected into E. coli
EPI300-T1® cells. Transformants were selected on Luria-Bertani (LB) agar containing
chloramphenicol (12.5 μgmL-1). To verify the size of the library fosmid DNA was extracted
from 24 clones and digested to completion with HindIII and EcoRI (Fermentas). Insert size
was estimated by agarose gel electrophoresis and visualization with an Alpha Imager 3400
imaging system.
Construction of 16S rRNA gene clone library and phylogenetic analysis
Transformants plated on LB agar were pooled and total fosmid DNA extracted using a
Qiagen®Midi kit. The 16S rRNA gene was amplified from the purified fosmid DNA using
the universal bacterial primers E9F and U1510R [10, 11]. Cycling conditions for the
universal bacterial primers included denaturation (4 min at 94 °C), followed by 30 cycles of
denaturation (30 s at 94 °C), annealing (30 s at 52 °C), and extension (105 s at 72 °C),
followed by a final extension (10 min at 72 °C). PCR was carried out in 50 μl reaction
volumes containing 2 mM MgCl2, 1 U Dream Taq polymerase (Thermo Scientific), 1 X PCR
buffer, 200 μM dNTPs, 0.5 μM of each primer and 100 ng fosmid DNA. PCR products were
purified with a GFXT PCR DNA purification kit (Illustra) and cloned into pGEM®-T Easy
vector (Promega) according to the manufacturer‟s instructions. Putative recombinant clones
were screened by colony PCR using universal M13 primers. Compost fosmid library (CFL)
clones were grouped manually into ribotypes based on ARDRA restriction patterns generated
by single digestions using AluI and BsuRI (isoschizomer of HaeIII) (Fermentas). Partial 16S
rRNA gene sequences were obtained for at least one representative clone from each ribotype.
Chromatograms were edited with Chromas software version [12] and sequences were
assembled in DNAMAN version 4.13 (LynnonBioSoft). Local alignments were obtained by
performing a standard nucleotide-nucleotide BLAST search (BLASTn) [13] of the GenBank
database. For phylogenetic analysis reference strains identified from the BLAST search were
selected for comparison. Sequences were aligned using CLUSTAL_W [14] and checked
manually for errors. Phylogenetic analyses were conducted using MEGA version 5.0 [15]
and trees were constructed using the neighbour-joining [16] algorithm. The robustness of
tree topology was evaluated by bootstrap analysis [17] based on 1,000 resamplings.
High-throughput screening of fosmid library clones
The library (approximately 105 clones) was inoculated into 384-well microtitre plates
containing 50 µl LB broth supplemented with 12.5 µg mL-1 chloramphenicol using a QPix2
automated colony picker (Genetix). Primary screening was performed by gridding the library
from the 384-well microtitre plates onto 22 x 22 cm Q-trays (Genetix) containing 500 ml
basal medium (LB agar) supplemented with 12.5 µg mL-1 chloramphenicol (Sigma) and
0.01% (w/v) L-arabinose (Sigma). Clones were screened for general esterase activity on basal
agar containing 1% (w/v) Gum Arabic (Sigma) and 0.1% (v/v) glycerol tributyrate (Sigma).
A clear halo surrounding a colony indicated lipolytic and/or esterase activity [18]. Positive
esterase clones were screened for ferulic acid esterase activity by plating onto basal agar
supplemented with 0.4% (w/v) ethyl 4-hydroxy-3 methoxycinnamate (ethyl ferulate)
(Sigma). A clear halo surrounding a colony indicated ferulic acid esterase activity [19].
Fosmid Pyrosequencing and ORF analysis
Equimolar concentrations of DNA from 36 fosmids were pooled and sequenced on a Roche
454 GS FLX sequencer by Inqaba Biotechnology, Pretoria, South Africa. Sequence reads
were assembled with CLC Genomics Workbench software (www.clcbio.com) and
Sequencher® sequencing software (www.genecodes.com) to produce contiguous sequences.
ORFs were predicted using Softberry‟s bacterial operon and gene prediction tool, FGENESB
(http://linux1.softberry.com) [20]. Signal peptides were predicted with SignalP version 4.0
[21]. Homology and similarity searches of the translated sequences of the predicted ORFs
were performed using the BLASTp program [13, 22, 23].
DNA sequence analysis
Multiple alignments of the predicted amino acid sequence of EstG34 were performed using
MEGA 6.0 [24]. A phylogenetic tree of EstG34 with other family VIII esterases, class-C
β-lactamases, ferulic acid esterases and DD-peptidases was constructed using the neighbourjoining method [16]. The robustness of tree topology was evaluated by bootstrap analysis
[17] based on 1,000 resamplings.
Expression and purification of recombinant EstG34
The EstG34 gene was amplified using fosmid 6-C1 as the template and the following
GAGGTC GAC GAT GGC CGA ATA-3‟ (nucleotides in bold denote NdeI and XhoI
restriction enzyme sites, respectively). The EstG34 gene was cloned into the expression
vector pET-21a (+) and the recombinant plasmid was transformed into E. coli Rosetta pLysS
cells. Cells were cultured to an optical density of approximately 0.8 at 600 nm at 30 °C, after
which 0.8 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) was added to the culture for
the induction of protein expression. Following 5 h of cultivation at 30 °C, cells were
harvested by centrifugation (6,000 x g for 15 min at 4 °C) and resuspended in 1X PBS buffer
(pH 7.4). Cells were disrupted by sonication and cellular debris was harvested by centrifuged
(8,000 x g for 15 min at 4 °C). EstG34 was purified by HIS-tag nickel affinity
chromatography using the HIS-Bind® resin and buffer kit (Novagen, USA) according to the
manufacturer‟s instructions. The eluate was dialysed overnight in a 6 ml Slide-A-Lyzer®
cassette (Thermo Fisher Scientific) against 50 mM sodium phosphate; 50 mM sodium
chloride (pH 7.9) buffer and stored at 4 °C. The protein concentration was estimated by the
method of Bradford [25] using the Bio-Rad protein assay kit with bovine serum albumin as a
standard. Protein purity was examined by sodium dodecyl sulphate-polyacrylamide gel
electrophoresis (SDS-PAGE) under denaturing conditions, as described by Laemmli [26].
Esterase activity assay
Esterase activity assays were performed using a standard colorimetric method by measuring
the release of p-nitrophenol from p-nitrophenyl (p-NP) esters. The release of p-nitrophenol
was monitored continuously at 410 nm using a Cary 50 Bio spectrophotometer (Varian, CA,
USA). All assays were prepared and analysed in triplicate. Enzyme activity was measured at
25 °C and one unit of activity was defined as the amount of enzyme releasing 1 µmol of
p-nitrophenol per minute under the defined assay conditions. The effect of pH on esterase
activity was studied by measuring activities on p-NP decanoate over a pH range of 4.5 – 11.
The following range of buffer systems were used: 100 mM sodium acetate (pH 4.5 to 5.5),
100 mM morpholineethanesulfonic acid (MES) (pH 5.5 to 7), 100 mM TRIS-HCl (pH 7 to
9), and 100 mM N-cyclohexyl-3-aminopropanesulfonic acid (CAPS) (pH 9 to 11). The
optimal temperature for enzyme activity was determined for a temperature range of 20 to
50 °C using the standard assay towards p-NP decanoate. To test thermal inactivation the
recombinant enzyme was preincubated at a range of temperatures for 1 hr. Residual esterase
activity was assayed on p-NP propanoate at 25 °C in 100 mM sodium phosphate pH 7.5, 100
mM sodium chloride.
Substrate specificity assays
p-nitrophenyl esters:
The hydrolytic activity of EstG34 against different fatty acid esters was investigated using the
following p-nitrophenyl esters: acetate (C2); propanoate (C3); octanoate (C8); decanoate
(C10); dodecanoate (C12) and hexadecanoate (C16).
Hydroxycinnamic acid methyl esters:
Activity towards the synthetic substrates methyl ferulate (Key Organics Ltd, Cornwall, UK),
methyl sinapate, methyl caffeate and methyl p-coumarate (APIN chemicals, Abingdon, Oxon,
UK) were based on the absorption difference of the free acid and the respective methyl ester.
The reaction was initiated by the addition of enzyme solution to the assay buffer (100 mM
sodium phosphate pH 7.5, 100 mM sodium chloride) containing 50 µM synthetic substrate
and was performed at 25 °C for 1 hour in a SPECTROstarNanomicroplate reader (BMG
Labtech). Absorbance was measured at 1 min intervals at 340 nm and activity was
determined using calibration curves of the substrate/product. Relative activities were
expressed as a percentage of the highest activity.
4-nitrophenyl ferulate:
Ferulic esterase activity of EstG34 was determined quantitatively using 4-nitrophenyl ferulate
(4-NPF) as the substrate [27]. The substrate was synthesized according to the method of
Hegde et al. [28]. The assay was carried out in 100 mM sodium phosphate (pH 7.5); 100 mM
sodium chloride buffer containing 1 mM 4-NPF. The liberated free p-nitrophenol was
measured at 410 nm. One unit of enzyme activity is defined was the amount of enzyme
releasing 1 µmol of p-nitrophenol from 4-NPF in 1 min at 25 °C.
Feruloyl esterase activity was also determined using the substrates 4-nitrophenyl 5-O-transferuloyl-α-L-arabinofuranoside (NPh-5-Fe-Araf) and 4-nitrophenyl 2-O-trans-feruloyl-α-Larabinofuranoside (NPh-2-Fe-Araf). These substrates were synthesised as detailed by
Mastihubova et al. [27]. The assay was carried out in 100 mM sodium phosphate buffer (pH
6.5) containing 12 l DMSO and 3 l Tween 20 per ml substrate (NPh-5-Fe-Araf or NPh-2Fe-Araf at 2.5 mM) [29]. The reaction mixture was incubated to 30 °C before addition of
EstG34. Measurements of absorbance at 420 nm were taken over a period of 3 hours and 1
unit of activity was defined as an absorbance change of 1.0 min-1 (A420)
Naphthyl acetate:
Deacetylase activity was determined by measuring α-naphthol released from naphthyl acetate
using the Fast Garnet liquid assay as described by Koseki et al. [30]. The assay was
performed in 50 mM sodium phosphate buffer (pH 7.5) with 0.8 μmol α-naphthyl acetate
(Fluka). After incubation at 37 °C for 10 min, the reaction was terminated by the addition of
110 μL Fast Garnet GBC (6 mg mL-1 stock solution in 10% (w/v) SDS) and the absorbance
measured at 560 nm. One unit (U) of acetyl xylan esterase (AXE) activity was defined as the
amount of enzyme required to produce 1 μmol of product per min under the defined assay
Acetyl xylan hydrolysis:
To determine hydrolytic activity of EstG34 on acetyl xylan, an endoxylanase-free 1%
solution of acetyl xylan in 0.1 M sodium phosphate buffer (pH 6.5) was prepared. The
reaction mixtures with varying amounts of the enzyme (5 - 50 l) were incubated overnight at
37 °C in sealed, thin-walled glass test tubes. Precipitation in the reaction mixtures was
visually determined [31].
Kinetic Measurements
Esterase activity kinetics were determined for four different substrates: p-nitrophenyl acetate,
p-nitrophenyl propionate, p-nitrophenyl ferulate and α-naphthyl acetate. The reactions
Table 1: Kinetic parameters for EstG34 on selected synthetic substrates.
KM (mM)
Specific activity
(U mg-1)
p-nitrophenyl acetate
0.011 ± 0.001 26.25 ± 30
19.70 ± 30
1790 ± 204
p-nitrophenyl propanoate 0.023 ± 0.004 151 ± 11
113.25 ± 90
4913 ± 287
p-nitrophenyl ferulate
0.213 ±0.017
10.34 ±0.70
7.49 ± 0.81
35.13 ± 4.24
α-naphthyl acetate
0.84 ± 0.09
1337 ± 56
969 ± 48
1154 ± 74
contained different substrate concentrations (Table 1) and the parameters (KM and kcat) were
calculated by Michaelis-Menten direct plots generated using Graphpad Prism® version 5.02
(La Jolla, CA).
Effects of inhibitors and detergents
The effect of various metal ions (CaCl2, CoCl2, FeSO4, MgSO4, MnSO4, NiSO4 and ZnSO4),
EDTA, organic solvents (ethanol, methanol, isopropanol, acetone, acetonitrile and DMSO)
and chaotropic agents (CTAB, SDS, Tween 20, and Triton X-100) on EstG34 activity was
determined. The enzyme was incubated in the presence of the inhibitor/detergent in 100 mM
sodium phosphate (pH 7.5), 100 mM sodium chloride for 30 min at 25 °C, and assayed for
enzyme activity. The activity of the enzyme without the addition of metal ions or no exposure
to solvents or detergents was defined as 100% activity.
Nucleotide sequences
The 16S rRNA gene sequences amplified from the library have been submitted to the
GenBank database under the Accession Nos. HQ694846.1–HQ694892.1.The nucleotide
sequence for EstG34 has been deposited in the GenBank database under the accession
number KJ937963.
The compost metagenomic library was comprised of 150,000 clones with an average insert
size of 30.67 kb. The estimated 4.65 x 109 bp of insert DNA represents a coverage equivalent
to approximately 1,500 bacterial genomes [32]. Analysis of the bacterial diversity captured
within the fosmid library identified 39 ribotypes, of which 9 ribotypes were represented by a
single clone. Phylotype richness estimators SACE and SChao1 predicted 56 and 60 ribotypes,
respectively (data not shown).
Phylogenetic analysis, as shown in Supplementary Figure S1, revealed that the majority of
library clones were related to γ-Proteobacteria (63%), α-Proteobacteria (20%),
Bacteroidetes (7%) and Firmicutes (4%). The library was dominated by clones (70%) that
were most similar to uncultured bacteria from environmental or clinical sources, which is
common to other soil metagenomes [33]. Previous studies have shown that compost is a rich
source of microorganisms possessing lignocellulose hydrolyzing abilities [34, 35]. In this
study, several of the ribotypes identified were related to known lignocellulolytic enzymeproducing strains, confirming that genomes encoding lignocellulolytic enzymes were well
represented in the library (denoted by a black triangles ( ) in Supplementary Figure S1).
Approximately 105 fosmid clones were functionally screened for general esterase activity on
glyceryl tributyrate, and a total of 25 clones were confirmed positive for lipolytic activity
(25:100,000), a hit rate of 0.025%. The reported hit rates for esterase clones from
metagenomic studies appear to vary significantly, with values ranging from 0.87% from
termite guts [18] to 0.001% from various environmental soils (1:60,000) [36]. The esterase
positive clones were further analysed for ferulic acid esterase activity by screening on ethyl
ferulate. Four of the 25 clones demonstrated the ability to degrade this substrate. One of these
clones, pCC1fos6-C1, showed strong feruloyl esterase activity and contained a 1,233-bp ORF
(EstG34) encoding a polypeptide of 410 amino acids. No signal peptide was identified.
EstG34 displayed 77% primary sequence identity to a predicted β-lactamase from
Phenylobacterium zucineum HLK1 (YP_002129844) (Figure 1). The closest characterised
relative was EstM-N2 (49% amino acid identity), a metagenome derived cold-active esterase
belonging to the family VIII esterase/lipase and class C β-lactamases [37]. Further sequence
Fig 1 Evolutionary relationship of Est34 in relation to family VIII esterases, class C β- lactamases, type C and D feruloyl esterases [52], group 10 feruloyl esterases [53] and pencillin-binding
proteins. The phylogenetic tree was generated using the neighbour-joining method (MEGA 6.0 software). All protein sequences were retrieved from GenBank, with the associated accession
numbers indicated in parentheses. β-lactamase activity is reported for all family VIII esterases [+: hydrolysis of the β-lactam amide bond; +*: deacetylation of the β-lactam ring; +?:
unknown if activity represents hydrolysis of the ester or amide bond; -: no β-lactamse activity]. The numbers at the nodes indicate the bootstrap percentages based on 1000 replicates. The
bar represents 0.2 nucleotide substitutions per nucleotide position.
analysis searches using the Lipase Engineering database [38] and the Arpigny and Jaeger
classification scheme [5] indicated that EstG34 was a family VIII carboxylesterase.
Unlike other microbial esterase families where the active site serine residue is typically
located within the G-X-S-X-G pentapeptide motif, the serine residue of family VIII esterases
is situated within the S-X-X-K motif and serves as the catalytic nucleophile [5]. The
alignment (Figure 2) indicated that EstG34 contained the S-X-X-K consensus sequence (SM-T-K75). This S-X-X-K motif is conserved within most of the β-lactamase superfamily
proteins, which includes penicillin binding proteins (PBPs), DD-peptidases and other family
VIII carboxylesterases [39, 40, 41]. However, two other highly conserved class C
β-lactamase motifs (Y-A-N) and (K-T/S-G) [42] were not found in EstG34 (Figure 2).
EstG34 was expressed in E. coli and purified to near homogeneity by nickel-chelation
chromatography (Figure 3). Using p-NP esters as a substrate, EstG34 was most active at a
temperature of 41 ˚C and at pH 9. EstG34 showed a limited tolerance to low pH‟s, exhibiting
only 50% of its maximum activity at pH 7.0, while being completely inactivated at pH 6.0
(Figure 4A). EstG34 can be characterized as being mildly alkaliphilic, with high activity at pH
9.0 and 70% of maximal activity at pH 10.0. The composting process involves different
temperature stages, therefore despite having prepared the metagenomic library from material
in the thermophilic stage it is not unexpected that the library contained DNA from mesophilic
organisms. Furthermore, as the compost material contained bovine manure DNA from enteric
organisms may be present in the library. This may account for the relatively low thermal
stability of EstG34 (Figure 4B), which retained 90% of its activity after one hour at 40 °C and
was completely deactivated after 30 minutes at 50 °C (Figure 4C). Divalent cations showed no
significant influence on enzyme activity (Figure S2). Of the chaotropic agents tested, only 1
mM SDS showed significant inhibition of EstG34 activity (Figure S3), while the enzyme was
stable in all the solvents tested (Figure S3).
Fig. 2. Multiple sequence alignment of EstG34 and related Family VIII carboxylesterases, class C ␤-lactamases, penicillin-binding proteins, and class C and D feruloyl
esterases. Family VIII esterases are represented by EstB (Burkholderia gladioli, AAF59826), and the metagenome-derived esterases EstC (ACH88047) Est22 (AGT17593),
EstCE1 (AAY90134), EstA3 (AAZ48934), Est2K (ACX51146), EstU1 (AFU54388), AFU54388 EstM-N1 (AEA07653) and EstM-N2 (HQ154133). Class C ␤-lactamases are represented by Lac-1 (Escherichia coli, AAA23441), Lac-2 (Enterobacter cloacae, P05364) and AmpC (Citrobacter freundii, ACV32310). Penicillin-binding proteins are represented
by PBP-1 (Streptomyces sp. R61, P15555) and PBP-2 (Bacillus cereus, CAA09676). Feruloyl esterases are represented by AspChloro (Acinetobacter sp. ADP1, AAL54855),
EstB-Fae10 (Corynebacterium aurimucosum, YP 002835738), PSPPH (Pseudomonas syringae, YP 274582), pFFaeD (Penicillium funiculosum, AJ312296) and Fes (Opituts terrae,
YP 001817235). The conserved S-X-X-K carboxylesterase family VIII motif, as well as the Y-A-N and K-T/S-G class C ␤-lactamase motifs are shown.
Fig 3 SDS-PAGE (12%) electropherogram showing different fractions collected for G34 purification by HIS-bind nickel affinity chromatography: lane M – molecular weight markers, lane 1
– flow through fraction, lane 2 – binding fraction, lane 3 – wash fraction and lane 4 – elute fraction. (B) Purified G34 recombinant protein following dialysis. Lanes 1 – 3 showing increased
loading of the protein.
Fig. 4. (A) Effect of pH on activity of EstG34 toward p-NP decanoate. Assay buffers used were 100 mM sodium acetate (pH 4.5 to 5.5), 100 mM MES (morpholineethanesulfonic
acid) (pH 5.5 to 7), 100 mM Tris-HCl (pH 7 to 9), and 100 mM CAPS (pH 9 to 11). (B) Effect of temperature on activity of EstG34 toward p-NP decanoate. (C) Thermal inactivation
profile of EstG34 at 40 ◦ C ( ), 45 ◦ C ( ) and 50 ◦ C ( ). Activity is expressed as a percentage of that at zero time in the standard assay (100 mM sodium phosphate, 100 mM
NaCl (pH 7.5), 25 ◦ C) toward p-NP propionate. (D) Activity of EstG34 at 25 ◦ C towardsp-NP esters of various chain lengths (C2, acetate; C3, propanonate; C8, octanoate; C10,
decanoate; C12, dodecanoate; and C16, hexadecanoate) in 1 mL reactions containing 100 mM sodium phosphate (pH 7.5), 100 mM NaCl, 1% acetonitrile, and 0.5 mM p-NP
ester substrate. (E) Ferulic acid esterase activity of G34 towards hydroxycinnamic acid methyl esters, represented as relative activity (% from the highest activity). Substrates
are ordered (left to right) by increasing degree of substitution. MpC: methyl p-coumarate, MC: methyl caffeate, MF: methyl ferulate, MS: methyl sinapinate.
The hydrolytic activity of EstG34 was determined with various p-NP fatty acid esters and
other ester compounds. EstG34 demonstrated high catalytic activity against short-chain fatty
acids (Figure 4D). Maximum activity was obtained with p-NP acetate (C3) with greatly
decreased activity toward longer chain fatty acids (>C8). Kinetic analysis of EstG34 activity,
however, suggests that p-NP propanoate is the preferred substrate. Although a 2-fold higher
affinity for p-NP acetate was observed (Table 1), the kcat for p-NP propanoate was 5.8-fold
higher than that of p-NP acetate, and a higher kcat/KMvalue was observed for p-NP
propanoate. The kinetics of EstG34 were also assessed on the acetylated substrate α-naphthyl
acetate. EstG34 demonstrated the highest specific activity and turnover rate on this substrate,
although catalysis was more efficient with p-NP acetate. The much higher KM for α-naphthyl
acetate suggests that the lower affinity for this substrate is due to steric hindrance by the
bulky biphenolic structure of α-naphthyl. EstG34 showed no deacetylase activity on acetyl
xylan as demonstrated by the absence of precipitation.
On hydroxycinnamic ester substrates, EstG34 demonstrated preference in the order of
> methyl ferulate > methyl p-coumarate > methyl caffeate > methyl sinapinate (Figure 8). Of
all the substrates tested methyl sinapinate contains the highest degree of substitution and
EstG34 had approximately 80% less activity on this substrate compared to methyl ferulate.
The lower activity on substrates with a higher degree of substitution can be attributed to
increased steric congestion around the ester group. Feruloyl esterase activity was quantitated
using the substrates p-NP ferulate, p-NP 5-O-trans-feruloyl-α-L-arabinofuranoside and p-NP
2-O-trans-feruloyl-α-L-arabinofuranoside. EstG34 had a specific activity of 10.3 U mg-1, 0.65
U mg-1 and 0.11 U mg-1 on these substrates, respectively (Supplementary Figure S4). While
there are relatively few characterised feruloyl esterases in the literature [43, 44, 45] EstG34
activity is comparable to two characterised fungal feruloyl esterases [46].
Carboxylester hydrolases are ubiquitous enzymes and the current esterase classification
scheme consists of eight families (families 1-VIII), based primarily on sequence similarity,
and less so on biochemical properties [5]. EstG34, identified by the functional screening of a
compost metagenomic library, grouped with the family VIII esterases. This is a poorly
characterised esterase family, and has previously included class C β-lactamases, penicillin
binding proteins, DD-peptidases and a range of carboxylesterases. All family VIII esterases
demonstrate sequence identity to β-lactamases and of the top 100 BLASTp hits for EstG34
96 were annotated as β-lactamases. As a result, family VIII esterases are typically tested for
their ability to hydrolyse a variety of β-lactam substrates (Table 2). The majority either lack
the activity or show negligible activity, despite the high sequence identity to the
β-lactamases, while others have been described as exhibiting “promiscuous β-lactamase
activity” [18, 21, 41, 47, 48, 49]. Consequently, it has been suggested that these esterases
have evolved from the class C β-lactamases, where some have maintained this remnant
activity, while others have lost the capability due to steric interference resulting from
structural evolution [18, 41, 50].
EstG34 displayed no detectable activity when tested against ampicillin (data not shown).
Interestingly, detailed selectivity assays conducted for Est22 [49] and EstB [40] demonstrated
that the enzymes selectively hydrolysed the ester bond of a number of cephalosporin-based
substrates, while leaving the amide bond of the β-lactam ring intact. This clearly
differentiates them from bona fide β-lactamases. EstU1, on the other hand, cleaves the amide
bond of first generation β-lactam antibiotics, although it is more efficient at hydrolysing short
chain esters [48]. Other family VIII esterases (EstM-N1, EstM-N2, EstC) have demonstrated
activity on nitrocefin, although it is not clear from the data presented whether this constitutes
deacetylation or amide bond hydrolysis [18, 37]. The differentiation of deacetylation or
Table 2: Comparison of the specific activities of known family VIII esterases.
Specific Activity
(U mg-1)
151.3 Nd
Additional Substrates
α-naphthyl acetate
methyl caffeate
methyl ferulate
methyl sinapinate
naphthol AS-acetate
naphthol AS-2-chloro-acetate
2-naphthol 2-chloro-propanoate
linalyl acetate
Compost metagenome
This study
Cephalosporin C
Burkholderia gladioli
Acidic leachate
Acidic leachate
Cephalosporin C
Drinking water
vinyl acetate
vinyl propanoate
vinyl butyrate
vinyl acetate
vinyl propanoate
vinyl butyrate
vinyl caproate
vinyl parylate
vinyl laurinate
Compost metagenome
Soil metagenome
Arctic soil metagenome
+ : hydrolysis of the β-lactam amide bond
+* :- deacetylation of the β-lactam
+? : unknown if activity represents hydrolysis of the ester or amide bond
- : no β-lactamase activity
7-ACA: 7-aminocephalosporanic acid
lactamase activity is a crucial determinant and a key differentiation between family VIII
carboxylesterases from the β-lactamases. It is clear from the phylogenetic tree (Figure 1) that
the true β-lactamases cluster separately from the family VIII carboxylesterases. However, the
resolution of primary sequence comparisons does not sufficiently demonstrate the functional
evolution (clustering based on deacetylation versus amide bond hydrolysis, Figure 1), which
is expected to have resulted from a few critical amino acid changes in the catalytic site, as
opposed to global structural modifications. Crystallisation and mutation studies of EstU and
other family VIII esterases which have no activity on β-lactams should elucidate the active
site changes which determine the differences in substrate specificity. Irrespective of β-lactam
hydrolysis capacity, esters are the preferred substrates for all family VIII carboxylesterases
(Table 2) and the β-lactams are poor substrates. Due to the clear functional distinction
between class C β-lactamases and the carboxylesterases in family VIII, and for classification
clarification purposes, we propose that this family be referred to as carboxylesterases and not
β-lactamase fold enzymes.
EstG34 hydrolysed a wide variety of model esters, including hydroxycinnamic acids,
ferulated p-nitrophenyl, and α-naphthyl acetate, showing considerable preference for the
acetylated ester compounds. To the best of our knowledge, EstG34 represents the first family
VIII carboxylesterase with feruloyl esterase activities. Specificity towards hydroxycinnamic
acids has previously been used to sub-classify ferulic acid esterases (FAEs) as type A, B, C or
D [51]. Interestingly, despite the fact that EstG34 demonstrated hydrolytic properties
indicative of type C and D feruloyl esterases, it showed no significant sequence identity to
type C and D FAEs (Figure 2). However, the A-D classification system is restricted to
sequence similarity and substrate specificity on four model substrates only, and few fungal
FAEs are included [51, 52]. Considering that FAEs belong to highly divergent protein
superfamilies, with each family having a different evolutionary pathway to the genesis of
FAE specificity, a more robust classification scheme has been developed. The novel
descriptor-based classification system is a more reliable scheme which groups functionally
related FAEs that have common properties [52]. This was developed using FAEs belonging
to all protein superfamilies and representing fungal, bacterial and plant origins, and classifies
FAEs into 12 clusters (FEF1-12), which can be further sub-grouped based on the
constellation and distance between the catalytic triad residues. Furthermore, its robustness is
applicable even to poorly characterised enzyme families. EstG34 was predicted to group in
FEF10, which does not include any of the type C or D FAEs. Furthermore, EstG34 does not
show sequence similarity to FEF10 representatives (Figure 2). All FEF1-10 representatives
contain the catalytic triad (Ser, His, Asp), as well as the consensus “nucleophilic elbow”
(GXSXG, the universally conserved pentapeptide in which the catalytic serine residue is
located) [5], although the positioning of these in the sequence varies greatly. This
complicates EstG34 classification using the descriptor-based system since these conserved
motifs, which EstG34 lacks, are a prerequisite for the classification using the FAE descriptorbased system [52]. Furthermore, feruloyl estersases are characteristically α/β-fold hydrolases,
another characteristic that EstG34 lacks. EstG34 is the first FAE to be described which lacks
the characteristics traditionally used to classify feruloyl esterases.
This presents a fascinating question regarding the general classification of feruloyl esterases.
Both in this study and others, phylogenetic clustering of carboxylesterases does not correlate
with substrate specificity [52]. EstG34 also demonstrates that novel bacterial FAEs adopting
different structural folds and conserved motifs are yet to be discovered. While a preference
for short to medium chain length p-NP esters remains the benchmark for classifying an
enzyme as a „true carboxylesterase‟, the results presented here suggest that a refinement of
the classification of feruloyl esterases, and particularly the family VIII esterases, is required.
Specifically, the FAE classification now needs to consider the Family VIII type esterases
which harbour the (Ser, Lys, Tyr) catalytic triad where the serine is located in the conserved
(SXXK) motif. Given that many carboxylesterases show broad substrate ranges, and that a
limited number of key amino acid substitutions in the active site can dramatically affect
substrate specificity, we suggest that classification of these enzymes by ‟substrate preference‟
may be misleading. Certainly, characterisation using a limited range of synthetic nitrophenylester substrates offers little information on either the in vivo substrate or the classification of
the enzyme. EstG34, with high sequence identity to class C β-lactamases and falling within
the current classification scheme of both family VIII esterases and feruloyl esterases,
emphasises the current classification dilemma. As more EstG34 homologues are discovered,
and more in depth substrate characterisations of family VIII esterases is conducted, its
classification will become clearer.
There now exists a number of reports of CAZy enzymes possessing more than one catalytic
domain: for example, an enzyme from Prevotella ruminicola 23 contains two carbohydrate
esterase domains which separately confer acetyl- and ferulic esterase activities [53]. Another
example, Clostridium thermocellum XynY [54], contains xylanase and feruloyl esterase
domains (clustering with Type D FAEs) [45]. However, based on preliminary modelling
analyses, the multifunctional capability of EstG34 appears to be catalysed by a single
catalytic domain, with no obvious carbohydrate binding domains (data not shown).
EstG34 displayed the ability to hydrolyse a wide variety of ester bonds found in natural
lignocellulosic polysaccharides. Ferulated polysaccharides are essential in directing cell wall
cross-linkages and serve as a defensive mechanism against invasion by plant pathogens [54].
The deconstruction of hemicellulose requires the cooperative effect of a number of different
enzymes and microorganisms have developed cell associated multi-protein complexes, such
as cellulosomes [56] and xylosomes [57], containing cellulases, xylanases and carbohydrate25
binding modules to achieve this. O-acetyl and methyl esterification products are the most
frequently occurring substitutions found in various plant cell wall polysaccharides [58],
where short xylo-oligosaccharides could be de-acetylated by non-specific acetyl esterases
[31]. As the activity of endoxylanases can be partially or completely hindered by the presence
of acetyl groups [59], EstG34 could serve to improve the accessibility of endoxylanases to
the xylan backbone of plant polysaccharides.
This work was funded by The National Research Foundation (NRF) and the Technology
Innovation Agency (TIA), South Africa.
The authors declare that they have no conflict of interest.
[1] R.I Amann, W. Ludwig, K.H. Schleifer Microbiol. Rev. 59 (1995) 143-169.
[2] C.J. Duan, L. Xian, G.C. Zhao, Y. Feng, H. Pang, X.L. Bai, J.L. Tang, Q.S. Ma, J.X. Feng
J. Appl. Microbiol. 107 (2009) 245–256.
[3] N. Ilmberger, D. Meske, J. Juergensen, M. Schulte. P. Barthen, U. Rabausch, A. Angelov,
M. Mientus, W. Liebl, R.A. Schmitz, W.R. Streit Appl. Environ. Microbiol. 95 (2012) 135146.
[4] M. Allgaier, A. Reddy, J.I. Park, N. Ivanova, P. D'haeseleer, S. Lowry, R. Sapra, T.C.
Hazen, B.A. Simmons, J.S. Van der Gheynst, P. Hugenholtz, PLoS ONE 5 (2010) e8812.
[5] J.L. Arpigny, K.-E. Jaeger, Biochem. J. 343 (1999) 177-183.
[6] R. Gupta, N. Gupta, P. Rathi, Appl. Microbiol. Biotechnol. 64 (2004) 763-781.
[7] M. Levisson, J. van der Oost, S.W.M. Kengen, Extremophiles 13 (2009) 567-581.
[8] U.T. Bornsheuer, FEMS Microbiol. Rev. 26 (2002) 73-81.
[9] J. Zhou, M.A. Bruns, J.M. Tiedje, Appl. Environ. Microbiol. 62 (1996) 316-322.
[10] A.-L. Reysenbach, N.R. Pace, in F.T. Robb, A.R. Place (Eds.), Archaea: A Laboratory
Manual – Thermophiles, Cold Spring Harbour Laboratory Press, New York, 1995, pp 101107.
[11] M.C. Hansen, T. Tolker-Neilson, M. Givskow, S. Molin, FEMS Microbiol. Ecol. 26
(1998) 141-149.
[12] T.A. Hall, Nucleic acid Symposium Series 41 (1999) 95-98.
[13] S.F. Altschul, T.L. Madden, A.A. Schaffer, J.H. Zhang, Z. Zhang, W. Miller, D.J.
Lipman, Nucleic Acids Res. 25 (1997) 3389-3402.
[14] J.D. Thompson, D.G. Higgins, T.J. Gibson, Nucleic Acids Res. 22 (1994) 4673-4680.
[15] S. Kumar, K. Tamura, M. Nei, Brief. Bioinform. 5 (2004) 150-163.
[16] N. Saito, M. Nei, Mol. Biol. Evol. 4 (1987) 406-425.
[17] J. Felsenstein, Evolution 39 (1985) 783–791.
[18] K. Rashamuse, V. Magomani, T. Ronneburg, D. Brady, Appl. Microbiol. Biotechnol. 83
(2009) 491-500.
[19] J. Donaghy, P.F. Kelly, A.M. McKay, Appl. Microbiol. Biotechnol. 50 (1998) 257–260.
[20] V. Solovyev, A. Salamov, R.W. Li (Ed.), Automatic Annotation of Microbial Genomes
and Metagenomic Sequences, Nova Science Publishers, 2011, pp. 61-78.
[21] T.N. Petersen, S. Brunak, G. von Heijne, H. Nielsen, Nature Methods 8 (2011) 785-786.
[22] S.F. Altschul, J. Theor. Biol. 138 (1989) 297-309.
[23] A. Marchler-Bauer, J.B. Anderson, P.F. Cherukuri, C. De Weese-Scott, L.Y. Geer, M.
Gwadz, S. He , D.I. Hurwitz, J.D. Jackson, Z. Ke, C.J. Lanczycki, C.A. Liebert, C. Liu, F.
Lu, G.H. Marchler, M. Mullokandov, B.A. Shoemaker, V. Simonvan, J.S. Song, P.A.
Thiessen, R.A. Yamashita, J.J. Yin, D. Zhang, S. Bryant, Nucleic Acids Res. 33 (2005):
doi: 10.1093/nar/gki069
[24] K. Tamura, D. Peterson, N. Peterson, G. Stecher, M. Nei, S. Kumar, Mol. Biol. Evol. 28
(2011) 2731-2739.
[25] M.M. Bradford, Anal Biochem 72 (1976) 248-254.
[26] U.K. Laemmli, Nature 15 (1970) 680-685.
[27] V. Mastihuba, L. Kremnicky, M. Mastihubová, J.L. Willett, G.L. Côté, Anal. Biochem.
309 (2002) 96-101.
[29] S. Hegde, P. Srinivas, G. Muralikrisha, Anal. Biochem. 387 (2009) 128-129.
[30] P. Biely, M. Mastihubová, W.H. van Zyl, B.A. Prior, Anal. Biochem. 311(2002) 68-75.
[31] T. Koseki, K. Takahashi, S. Fushinobu, H. Iefuji, K. Iwano, K. Hashizume, H.
Matsuzawa, Biochim. Biophys. Acta. 1722 (2005) 200-208.
[32] K. Poutanen, M. Sundberg, H. Korte, J. Puls, Appl. Microbiol. Biotechnol. 33 (1990)
[33] E.M. Gabor, W.B.L. Alkema, D.B. Janssen, Environ. Microbiol. 6 (2004) 879-886.
[34] S.J. Joseph, P. Hugenholtz, P. Sangwan, C.A. Osborne, P.H. Janssen, Appl. Environ.
Microbiol. 69 (2003) 7210-7215.
[35] M.C. Vargas-Garcia, F. Suárez-Estrella, M.J. Lopez, J. Morena, Int. Biodeterior.
Biodegradation. 59 (2007) 322-328.
[36] C.M. Wang, C.L. Shyu, S.P. Ho, S.H. Chiou, Lett. Appl. Microbiol. 47 (2008) 46-53.
[37] Y.-J. Kim, G.-S. Choi, S.-B. Kim, G.-S. Yoon, Y.-S. Kim, Y.-W, Protein Expr. Purif. 45
(2006) 315-323.
[38] E.Y. Yu, M.-A. Kwon, M. Lee, J.Y. Oh, J.-E. Choi, J.Y. Lee, B.-K. Song, D.-H. Hahm,
J.K. Song, Appl. Environ. Microbiol. 90 (2011) 573-581.
[39] J. Pleiss, M. Fischer, M. Peiker, C. Thiele, R.D. Schmid, J. Mol. Catal. B: Enzym 10
(2000) 491-508.
[40] J.A. Kelly, O. Dideberg, P. Charlier, J.P. Wery, M. Libert, P.C. Moews, J.R. Knox, C.
Duez, C. Fraipont, B. Joris, J. Dusart, J.M. Frere, J.M. Ghuysen, Science 231 (1986) 1429–
[41] E.I. Petersen, G. Valinger, B. Solkner, G. Stubenrauch, H. Schwab, J. Biotechnol. 89
(2001) 11–25.
[42] Y.H. Kim, E.J. Kwon, S.K. Kim, Y.S. Jeong, J. Kim, H.D. Yun, H. Kim, Biochem.
Biophys. Res. Commun. 393 (2010) 45-49.
[43] B. Joris, J.M. Ghuysens, G. Dive, A. Renard, O. Dideberg, P. Charlier, J.M. Frere, J.A.
Kelly, J.C. Boyington, P.C. Moews, J. Biochem. 250 (1988) 313 – 324.
[44] M. Estaban-Torres, I. Reverón, J.M. Mancheño, B. de las Rivas, R. Muñoz, Appl.
Environ. Microbiol. 79 (2013) 5130-5136.
[45] D.W.S. Wong, V.J. Chan, H. Liao, M.J. Zidwick, J. Ind. Microbiol. Biotechnol. 40
(2013) 287-295.
[46] K. Rashamuse, T. Ronneburg, W. Sanyika, K. Mathiba, E. Mmutlane, D. Brady Appl.
Microbiol. Biotechnol. 98 (2014) 727-737.
[47] S. Hegde, G. Muralikrishna, World J. Microb. Biot. 25 (2009) 1963-1969.
[48] C. Elend, C. Schmeisser, C. Leggewie, P. Babiak, J.D. Carballeira, H.L. Steele, J.-L.
Reymond, K.-E. Jaeger, W. Streit, Appl. Environ. Microbiol. 72 (2006) 3637-3645.
[49] J.H. Jeon, S.-J. Kim, Y.S. Lee, S.-S. Cha, J.H. Lee, S.-H. Yoon, B.-S. Koo, C.-M. Lee,
S.H. Choi, S.H. Lee, S.G. Kang, J.-H. Lee, Appl. Environ. Microbiol. 77 (2011) 7830-7836.
[50] N. Mokoena, K. Mathiba, T. Tsekoa, P. Steenkamp, K. Rashamuse, Biochem. Biophys.
Res. Comm. 437 (2013) 342-348.
[51] U.G. Wagner, E.I. Petersen, H. Schwab, C. Kratky, Protein Sci. 11 (2002) 467-478.
[52] V.F. Crepin, C.B. Faulds, I.F. Connerton, Appl. Microbiol. Biotechnol. 63 (2004) 647652.
[53] D.B. Udatha, I. Kouskoumvekaki, L. Olsson, G. Panagiotou, Biotechnol. Adv. 29 (2011)
[54] M.A. Kabel , C.J. Yeoman , Y. Han , D. Dodd , C.A. Abbas , J.A. de Bont , M.
Morrison, I.K. Cann , R.I. Mackie, Appl. Environ. Microbiol. 77 (2011) 5671-5681.
[55] Blum
[56] N.D. Mohne, M. Bar-Peled, C. Somerville. In: M.E. Himmerl (Ed.) Biomass
recalcitrance, Blackwell, Oxford, 2008, pp 266 – 277.
[57] R. Lamed, E.A. Bayer, Adv. Appl. Microbiol. 33 (1988) 1-46.
[58] L.L. Lin, J.A. Thomson, FEMS Microbiol. Lett. 84 (1991) 197–204.
[59] P.M.-A. Pawar, S. Koutaniemi, M. Tenkanen, E.J. Mellerowicz, Front. Plant. Sci. 4
(2013) 1-8.
[60] P. Biely, C.R. Mackenzie, J. Puls, H. Schneider, Nature Biotechnology 4 (1986) 731–
Appendix A. Supplementary data
Supplementary Fig S1 Unrooted 16S rRNA gene phylogenetic tree of the compost fosmid library (CFL) clones
generated with general eubacterial primers. The tree was based on 615 bp of common sequence and was
obtained using the neighbour-joining method. Strains that produce lignocellulolytic enzymes are indicated with
by a black triangle ( ). GenBank accession numbers are given in parentheses.
Supplementary Fig S2 Effect of divalent cations and EDTA on the activity of G34. Relative activities are
provided as a percentage of the activity in the absence of cation.
Supplementary Fig S3 Effect of various denaturants, inhibitors and organic solvents on the activity of G34.
Supplementary Fig S4 Activity of G34 at 25°C towards a) 4-nitrophenyl 5-O-trans-feruloyl-α-Larabinofuranoside (NPh-5-Fe-Araf) and b) 4-nitrophenyl 2-O-trans-feruloyl-α-L-arabinofuranoside (NPh-2-FeAraf) in 100 mM sodium phosphate buffer (pH 6.5) Measurements of absorbance at 420 nm were taken over a
period of 3 hours and 1 unit of activity was defined as an absorbance change of 1.0 min-1 (A420).
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